Shifts in the relative abundance of ammonia-oxidizing bacteria and archaea in the water column of a stratified marine environment, Salt Pond

Sarah Hurley July 27, 2011 Department of Earth and Planetary Sciences, Harvard University 2011 Microbial Diversity Course, MBL

Abstract: Salt Pond, a seasonally stratified kettle pond in Falmouth, Massachusetts, provides an ideal analogue for the and other stratified marine environments. Nitrification in stratified water columns and minimum zones (OMZs) connects organic matter remineralization to the anammox and denitrification, the two known mechanisms of nitrogen loss in marine systems. We determined the effect of physiochemical gradients through the chemocline in Salt Pond on ammonia-oxidizing archaea and bacteria using amoA, the functional gene encoding ammonia monooxygenase subunit A. Ammonia oxidizing archaea (AOA) dominated the fresh, oxic surface water with low ammonium concentrations. Ammonia- oxidizing bacteria (AOB) abundance increased through the chemocline into the euxinic, but potentially micoraerophilic deep water. Within the AOA community, sequences were most related to cultured and uncultured putative AOA and grouped into surface, shallow, and deep clusters. The results of this study offer insights into the of AOA and AOB by comparing the environmental conditions in which these groups are numerically dominant. Introduction Nitrogen is an essential and often limiting nutrient fundamental to the structure and biochemistry of life. Nitrogen is readily oxidized or reduced and serves as both an electron donor and accepter in energy metabolisms of microorganisms. Microbial metabolisms drive the formation and consumption of different chemical form of nitrogen and therefore drive marine nitrogen biogeochemical cycling. Nitrification, the stepwise oxidation of ammonium to nitrite and nitrate, is a key process the marine nitrogen cycle responsible for the formation of the large deep-sea nitrate reservoir (Figure 1). Nitrification is a microbially mediated process, performed mostly by chemolithotrophic microbes who glean energy and electrons from this oxidation. Ammonia released from the remineralization of particulate organic nitrogen (PON or PN) is oxidized to substrates for denitrification and anammox. Nitrification therefore connects the recycling of organic nitrogen to the only two presently known nitrogen loss processes. In productive equatorial and coastal upwelling zones, high fluxes of organic matter and remineralization create oxygen minimum zones (OMZs), enabling denitrification (Hogslund et al., 2008) and anammox (Hamersley et al., 2007). Remineralization releases large amounts of ammonium, driving the high nitrification rates often associated with OMZs. Nitrogen loss from OMZs are estimated to account for 30-50% of total nitrogen loss from the (Gruber & Sarmiento, 1997). The seasonally meromictic kettle pond, Salt Pond, in Falmouth Massachusetts provides an ideal model system to study nitrogen cycling processes through an oxygen gradient. The water column in Salt Pond closely resembles the chemical and oxygen gradients typically associated with large marine anoxic basins such as the Black Sea (Damste et al., 1993). The water column in these systems consists of an oxic surface layer, sulfidic and anoxic deep waters, and consistent concentrations of chemical species along through the transition zone, or chemocline. Studies of nitrification processes, such as ammonia oxidation, in Salt Pond can there be translated to OMZs and larger stratified marine systems. Ammonia oxidation, the first step in nitrification, is considered rate-limiting and nitrite rarely accumulates in natural environments (Schleper & Nicol, 2010). Traditionally, ammonia oxidation was thought to be catalyzed by Nitrosomona and Nitrospira in the betaproteobacteria (Head et al., 1993), and Nitrosococcus in the gammaproteobacteria (Purkhold et al., 2000; Ward & O'Mullan, 2002). Metagenomic studies provided the first evidence for the metabolic capacity to oxidize ammonia in marine group 1 Crenarchaeota (Venter 2004, Treusch). The first successful isolation of an autotrophic ammonia-oxidizing marine archaeon, Nitrosopumilus maritiums (Könneke et al., 2005), led to the discovery that mesophilic ammonia-oxidizing archaea (AOA) are ubiquitous in the global oceans (Francis et al., 2005b). In the open and suboxic basins, AOA outnumber AOB in the water column (Coolen et al., 2007; Lam et al., 2007). The first step in ammonia oxidation, the oxidation of ammonia to hydroxylamine, is catalyzed by the membrane bound ammonia monooxygenase (AMO) enzyme (Könneke et al., 2005). AMO is encoded by the amoA, amoB, and amoC genes. Putative crenarchaeal homologues to all three subunits have been identified (Hallam et al., 2006; Treusch et al., 2005). Due to its functional significance and conserved phylogeny, the amoA gene can be used as a molecular marker to study the diversity and abundance of AOA and AOB (Coolen et al., 2007; Lam et al., 2007). This study attempts to address the following questions on the abundance and diversity of AOA and AOB in the stratified, marine analog Salt Pond: 1. Who is the ammonia oxidizing community? 2. Where does this community occur in the water column? 3. How to physiochemical parameters affect the distribution of ammonia oxidizing archaea vs. ammonia oxidizing bacteria? The following approaches were taken to address these questions: 1. 454 16S sequncing and community analysis at 9 depths 2. Functional gene (amoA) clone libraries 3. qPCR targeting archaeal and bacterial amoA 4. CARD-FISH and DAPI cell counts

Materials and Methods Site description: Salt Pond is a brackish, shallow, seasonally stratified pond located in Falmouth, Massachusetts (Figure 2). It is a glacially formed kettle pond ranging 4.6 to 5.8 meters deep, with a surface area of 0.29 km2 (Simmons et al., 2004), and is surrounded by 0.02 km2 of upstream salt marsh (http://www.capecodcommission.org/tidalatlas/). The pond exhibits density due to permanent tidal exchange with Vineyard Sound and freshwater inputs from runoff, precipitation, and groundwater. The average daily tidal amplitude is ~0.6 m.

Sampling: Sampling was conducted from an inflatable rowboat on July 8, 2011. A YSI 600R Water Quality Sampling Sonde (Yellow Springs, Inc., Yellow Springs, Ohio) was used to collect (ppt), (°C), oxygen (% DO, mg/L) and pH data. A profile was taken in two locations ~10 meters apart to determine the water depth and location of the chemocline. The YSI probe was then attached to the intake of a portable peristaltic pump to allow for simultaneous water collection and profiling (Figure 3). Water samples were collected from 9 depths spanning 0-4.0m. Water samples for molecular work and FISH analysis were collected in acid rinsed 2L polycarbonate bottles. In an attempt to measure sulfide concentrations 1 ml of water was taken from the full 2 L with a tipless syringe and transferred into 25 ml of zinc acetate solution (25 ml Milli-Q water + 1 ml 20% by volume zinc acetate). This procedure did not adequately fix the sulfide or was too large a dilution as no sulfide was detected using the Cline assay (Cline, 1969).

FISH: Immediately on return to lab, water samples for FISH analysis were fixed using 1% formaldehyde in 50 ml conical tubes. 2 and 5 ml aliquots were filtered onto 0.2 µm membrane filters (Millipore GTTP, 25 mm). CARD-FISH was performed using the following probes:

Eub338 Bacterial Eub338 II Eub338 III Cren537 Crenarchaeal Cren554 Eury806

Nutrient Analyses: 50 ml of water was sterile filtered and frozen for nutrient analysis. Ammonia measurements were performed by Chris Algar at the MBL Ecosystem Center according to (Scheiner, 1976). Nitrite measurements were performed spectrophotometrically according to (Strickland J, 1968).

Community 16s RNA analysis: Between 200-300 ml of water from each sampling depth was filtered onto 0.2 µm membrane filters (Millipore GTTP, 47 mm) for DNA extraction. DNA for 454 sequencing was extracted using a PowerSoil DNA Isolation Kit and quantified using a NanoDrop spectrophotometer (Thermo Scientific). SSU rRNA genes were amplified with barcoded primers that also incorporated the Roche 454 Ti adapter sequences. The barcode was on the forward primer. Each barcode was 9 nt long and all the barcodes we used are in the mapping file for the 454 data.The primer targets are 515F and 907R on the E. Coli 16S gene.

Forward primer (X denotes barcode, lowercase is the linker between barcode and 16S primer), 5'-CGTATCGCCTCCCTCGCGCCATCAGXXXXXXXXgaGTGYCAGCMGCCGCGGTAA-3'

Reverse primer (lowercase denotes linker between adapter and 16S primer), 5'-CTATGCGCCTTGCCAGCCCGCTCAGggCCGYCAATTCMTTTRAGTTT-3' In the "AmplificationPlate" in the mapping file "plate1" was amplified for 32 cycles and "plate4" in the "AmplificationPlate" field was amplified for 36 cycles. The PCR program utilized a touchdown annealing temp for the first 10 cycles from 68-58C. Then there were 12 cycles of three-step PCR (denaturation, annealing, elongation) followed y 10-14 cycles of tow-step PCR (annealing and elongation at same temp). A Phusion HF polymerase (2X mastermix) was used to amplify the gene with 8% DMSO and 0.5 uM primers in the final reaction volume. The template was normalized to 15 ng/uL (for DNA above 15 ng/uL) and 2 uL of each template was used forPCR. Post amplification DNA was quantified using the PicoGreen assay and then pooled ~125 ng of each PCR product. This pool was concentrated down to 100 uL using the vacufuge and gel purifed using the Montage Kit (Millipore). The gel- purified pool was then shipped to Penn St. for sequencing. The RDP classifier as implemented in the Qiime pipeline for community microbial analysis tool was used to assign taxa to all sequences. Trees were built using the GreenGenes NAST aligner tool to align sequnces with the core set of 16s alignment templates. Sequences were aligned using the filter_alignment.py protocol in Qiime with lane masking enabled (as described and referenced by Libusha Kelly, course report, 2010).

Clone Libraries and qPCR: DNA for all other molecular analyses was extracted using a phenol- chloroform, freeze-thaw procedure in the Coolen Lab at WHOI. Total nucleic acid extract was with a ladder in a 1% agarose and visualized with ethidium bromide (Figure 4). DNA extracts were quantified using fluorometry and yielded the following concentrations:

ng/µl Sample Depth DNA SP-A 0.0 m 178.5 SP-B 0.5 m 309.6 SP-C 1.0 m 239.6 SP-D 1.5 m 12.5 SP-E 2.0 m 168.3 SP-F 2.5 m 148.9 SP-G 3.0 m 121.6 SP-H 3.5 m 62.2 SP-I 4.0 m 116.7

PCR for archaeal and bacterial amoA clone libraries and qPCR were performed using primers and thermocycling conditions previously described for bacterial amoA (AmoA-1F*/AmoA-2R) specifically targeting the β-AOB (Rotthauwe et al., 1997), and archaeal amoA (Arch-amoAF/Arch-amoAR) (Beman et al., 2008; Francis et al., 2005a). An inhibition test was performed to check for inhibition due to remaining extraction solvents, etc. Significant inhibition was not demonstrated in any of the samples. Clone libraries using archaeal primers as described above were constructed for depths 0 m, 0.5 m, 1.5 m, and 4.0 m. PCR products were cloned into the TOPO2 TA Cloning Kit (Invitrogen). Sequences in the clone libraries were aligned against each other using Muscle. Sequences in each library were locally BLASTed against the a FunGene amoA database. A tree was constructed using FastTree and visualized in iTOL.

Results and Discussion: Salt Pond Depth Profile: Seasonal density stratification in Salt Pond occurs during the summer months– late June through September– and prevents vertical mixing (Simmons et al., 2004). Sampling on July 8 revealed a distinct chemocline from 1.5 to 3 meters (Figure 5), as reported in previous years (Saenz, 2008). The surface of the pond is oxygenated freshwater with a salinity of 11.37 ppt, dissolved oxygen concentration of 6.4-6.7, and a pH of 8.8. Respiratory oxygen consumption drives down dissolved oxygen that cannot be replenished by mixing and creates a chemocline. Below the chemocline the salinity increases to 19 ppt, the dissolved oxygen drops to 4.1 mg/L, and the pH decreases to 7. Sulfide concentrations likely increase to below the oxycline to millimolar concentrations (Saenz, 2008).

16S Community Analysis by 454 sequencing: The major phyla represented by 16S rRNA 454 sequencing are Actinobacteria, Bacteroidetes, Chlorobi, Cyanobacteria, and Proteobacteria. Cyanobacteria are the dominant phyla in water column down to 2.5 meters with the exception of 0.5m where Proteobacteria (Figure 6). While photosynthetically active radiation (PAR) was not measured, this abundance pattern reflects photosynthetic activity in Cyanobacteria and Proteobacteria through the . At 3 meters there is a slight relative increase in the abundance of Bacteroidetes followed by a complete dominance of Chlorobi (97% of relative phyla counts at 4 meters). The increase in Chlorobi at 3.5-4 meters reveals the spatial niche of low-light adapted green sulfur bacteria in the euxinic bottom waters of Salt Pond. The time constraints of this course did not allow for a very in-depth analysis of this dataset. For example, breaking down the proteobacteria could provide information on the spatial niche and relative representation of and ammonia oxidizing bacteria. In order to assess the difference in microbial communities at each depth, weighted and unweighted dendrograms were produced using UniFrac (Figure 7). The UniFrac method measures the phylogenetic distance between sets of taxa in a phylogenetic tree as the fraction of the branch length of the tree that leads to individual descendants from each water column depth (Lozupone & Knight, 2005). Given the gradient of physiochemical conditions in Salt Pond, a logical hypothesis would be that microbial communities at each depth would be the similar to the communities at adjacent water depths. To a coarse degree this appears to be true in the dendrograms. Samples, especially in the more closely chemically related surface and deep water tend to cluster by depth, while samples in the more rapidly changing chemocline are more spread out. In this case the 0.5 m sample may be reflecting a photosynthetic community different from the otherwise dominant cyanobacteria. Rarefaction curves for the 16S community analysis do not show saturation at any depth (Figure 8). This is unsurprising as each depth was only sampled once. More notably, the grouping of the rarefaction curves mirrors patterns found in the relative representation of phyla and the UniFrac dendograms, largely that the sample from 0.5 meters was the only sample approaching a plateau. qPCR: qPCR was performed on the total nucleic acid extracts at each depth to look at the relative abundance of putative ammonia-oxidizing archaea (AOA) and bacteria (AOB) across the physiochemical gradient of the chemocline in Salt Pond. Shifts in the relative abundance of AOA and AOB correlate closely with both dissolved oxygen and ammonium concentration through the water column. In the first ½ meter of the water column AOB dominate ~70% of amoA community. Between ½ and 2 meters, in the oxic to suboxic water just above the chemocline, AOA make up over 95% of the amoA community. Through the chemocline and into the anoxic deep water, the abundance of AOB increases steadily to 66%. In the lower 3 meters of the water column the increase in the abundance of the AOB community covaries with the increase in ammonium concentration. In the surface water of Salt Pond, the high percentage of AOB may be due to nutrient run off of surface water. In past years, high ammonium concentrations have been observed in the upper meter of Salt Pond, so this may be a temporal variation that was missed on the one sampling expedition. The location of a peak abundance of AOA in the oxic to suboxic nitrification zone just above the chemocline has been similarly observed in the Black Sea (Lam et al., 2007) associated with maximum nitrification activity. In and below the chemocline with the decrease of dissolved oxygen, the remineralization of organic matter and grodundwater inputs provide a source of ammonium. As ammonium concentrations increase and dissolved oxygen concentrations decrease, AOB become more prevalent than AOA. This suggests that AOB outcompete AOA at high ammonia concentrations in a potentially microaerophilic environment created by an influx of ground water from the sediments. Functional Gene Clone Libraries: Clone libraries targeting the archaeal amoA gene were locally BLASTed against the FunGene amoA database. While the results were not particularly illuminating, they were the similar to uncultured and enrichment cultures of ammonia oxidizing archaea and Crenarchaeota (Table 1). Trees constructed from archaeal amoA clone libraries showed roughly three clustering groups (Figure 10). There is a ‘very shallow’ depth (0-0.5m) cluster, a ‘shallow’ depth cluster, and a very distinct ‘deep’ cluster. This pattern appears in both the tree ignoring branch lengths and the tree incorporating branch lengths. In the tree incorporating branch lengths, the ‘deep’ clustering is more readily apparent as the branch length of the cluster is distinct from both the ‘very shallow ‘ and ‘shallow’ clusters on either side.

CARD-FISH and DAPI Counts: CARD-FISH counts with crenarchaeal and general bacterial probes revealed a consistent ratio of bacteria to crenarchaea throughout the water column. The crenarchaea generally make up ~1% of the total DAPI counts and the bacteria make up between 70-80% of the counts. 20-30% of the DAPI counts were not fluorescently labeled with either probe. Morphology in the crenarchaeal probe resembled the peanut shape observed in (Könneke et al., 2005).

Conclusions: • Ammonia-oxidizing crenarchaea and bacteria are both present in Salt Pond. AOA are dominant in the oxic surface waters with low ammonia concentrations while AOB increase through the chemocline and into the suboxic, ammonia rich waters • The putative ammonia-oxidizing archaea are closely related to cultured and uncultured environmental ammonia-oxidizers in the FunGene amoA database. • CARD-FISH revealed a low overall abundance of crenarchaea compared to bacteria throughout the water column. • The microbial community changes significantly through water column in Salt Pond, highlighting different ecological niches in a stratified marine system.

Future Directions and Remaining Questions: • Phylogenetic comparison to known ammonia-oxidizing organisms: Given the lack of specificity in the BLAST results from the FunGene amoA database, the next step would be to build a tree with the clone libraries from Salt Pond to compare to known organisms of interest such as Nitrosopumilus maritiums and Crenarchaeota found in the Black Sea (Coolen et al., 2007). • Nitrification rates: Rate measurements for ammonia oxidation and nitrite oxidation would allow us to look at where in the water column, nitrification is occurring relative to the numerical abundance of either AOA or AOB. • Temporal variation: Salt Pond is a seasonally stratified pond. The chemocline develops around late June, so sampling on July 8, 2011 is representative of an early summer chemocline. The ammonia-oxidizing community may shift with the evolution of the chemocline throughout the summer, and likely redistributes with the vertical mixing in winter. • Anammox: Below the chemocline Salt Pond becomes euxinic and could potentially support an active anammox community. This could be accomplished through similar methods used in this report or ladderane lipids unique to anammox bacteria. • Further 16S analysis: The 454 sequencing provided a depth of data, which could not be fully explored during the limited time in this course. Further probing of community structure at each depth would likely further our understanding of spatial niches in the stratified marine environment of Salt Pond.

Acknowledgements: Thank you to course directors Dan Buckley and Steve Zinder for providing this truly unique and fulfilling opportunity. Thank you to the course TAs (especially Chuck and Lizzy) for your help and patience. Thank you to Cornelia Wuchter and Marco Coolen for primers, probes, lab space, and scientific guidance. Thank you to the waterbears for rocking the hole.

Funding provided by the Milston L. Shifman Endowed Scholarship.

References: Beman, J.M., Popp, B.N., Francis, C.A., (2008) Molecular and biogeochemical evidence for ammonia oxidation by marine Crenarchaeota in the Gulf of California. Isme Journal, 2(4), 429-441. Cline, J.D., (1969) SPECTROPHOTOMETRIC DETERMINATION OF HYDROGEN SULFIDE IN NATURAL WATERS. Limnology and Oceanography, 14(3), 454-&. Coolen, M.J.L., Abbas, B., van Bleijswijk, J., Hopmans, E.C., Kuypers, M.M.M., Wakeham, S.G., Damste, J.S.S., (2007) Putative ammonia-oxidizing Crenarchaeota in suboxic waters of the Black Sea: a basin-wide ecological study using 16S ribosomal and functional genes and membrane lipids. Environmental Microbiology, 9(4), 1001-1016. Damste, J.S.S., Wakeham, S.G., Kohnen, M.E.L., Hayes, J.M., Deleeuw, J.W., (1993) A 6,000-YEAR SEDIMENTARY MOLECULAR RECORD OF CHEMOCLINE EXCURSIONS IN THE BLACK-SEA. Nature, 362(6423), 827-829. Francis, C., Roberts, K., Beman…, J., (2005a) Ubiquity and diversity of ammonia- oxidizing archaea in water columns and sediments of the ocean. In: Proceedings of the …. Francis, C.A., Beman, J.M., Kuypers, M.M.M., (2007) New processes and players in the nitrogen cycle: the microbial ecology of anaerobic and archaeal ammonia oxidation. Isme Journal, 1(1), 19-27. Francis, C.A., Roberts, K.J., Beman, J.M., Santoro, A.E., Oakley, B.B., (2005b) Ubiquity and diversity of ammonia-oxidizing archaea in water columns and sediments of the ocean. Proceedings of the National Academy of Sciences of the United States of America, 102(41), 14683-14688. Gruber, N., Sarmiento, J.L., (1997) Global patterns of marine nitrogen fixation and denitrification. Global Biogeochemical Cycles, 11(2), 235-266. Hallam, S.J., Mincer, T.J., Schleper, C., Preston, C.M., Roberts, K., Richardson, P.M., DeLong, E.F., (2006) Pathways of carbon assimilation and ammonia oxidation suggested by environmental genomic analyses of marine Crenarchaeota. Plos Biology, 4(4), 520-536. Hamersley, M.R., Lavik, G., Woebken, D., Rattray, J.E., Lam, P., Hopmans, E.C., Sinninghe Damste, J.S., Krueger, S., Graco, M., Gutierrez, D., Kuypers, M.M.M., (2007) Anaerobic ammonium oxidation in the Peruvian oxygen minimum zone. Limnology and Oceanography, 52(3), 923-933. Head, I.M., Hiorns, W.D., Embley, T.M., McCarthy, A.J., Saunders, J.R., (1993) THE PHYLOGENY OF AUTOTROPHIC AMMONIA-OXIDIZING BACTERIA AS DETERMINED BY ANALYSIS OF 16S RIBOSOMAL-RNA GENE-SEQUENCES. Journal of General Microbiology, 139, 1147-1153. Hogslund, S., Revsbech, N.P., Cedhagen, T., Nielsen, L.P., Gallardo, V.A., (2008) Denitrification, nitrate turnover, and aerobic respiration by benthic foraminiferans in the oxygen minimum zone off Chile. Journal of Experimental Marine Biology and Ecology, 359(2), 85-91. Könneke, M., Bernhard, A., José, R., Walker, C., (2005) Isolation of an autotrophic ammonia-oxidizing marine archaeon. In: Nature. Lam, P., Jensen, M.M., Lavik, G., McGinnis, D.F., Mueller, B., Schubert, C.J., Amann, R., Thamdrup, B., Kuypers, M.M.M., (2007) Linking crenarchaeal and bacterial nitrification to anammox in the Black Sea. Proceedings of the National Academy of Sciences of the United States of America, 104(17), 7104-7109. Lozupone, C., Knight, R., (2005) UniFrac: a new phylogenetic method for comparing microbial communities. Applied and Environmental Microbiology, 71(12), 8228-8235. Purkhold, U., Pommerening-Roser, A., Juretschko, S., Schmid, M.C., Koops, H.P., Wagner, M., (2000) Phylogeny of all recognized species of ammonia oxidizers based on comparative 16S rRNA and amoA sequence analysis: Implications for molecular diversity surveys. Applied and Environmental Microbiology, 66(12), 5368-5382. Rotthauwe, J.H., Witzel, K.P., Liesack, W., (1997) The ammonia monooxygenase structural gene amoA as a functional marker: Molecular fine-scale analysis of natural ammonia-oxidizing populations. Applied and Environmental Microbiology, 63(12), 4704-4712. Scheiner, D., (1976) DETERMINATION OF AMMONIA AND KJELDAHL NITROGEN BY INDOPHENOL METHOD. Water Research, 10(1), 31-36. Schleper, C., Nicol, G.W., (2010) Ammonia-Oxidising Archaea - Physiology, Ecology and Evolution. In: R.K. Poole (Ed.), Advances in Microbial Physiology, Vol 57, 57, Advances in Microbial Physiology (Ed. by R.K. Poole), pp. 1-41. Simmons, S.L., Sievert, S.M., Frankel, R.B., Bazylinski, D.A., Edwards, K.J., (2004) Spatiotemporal distribution of marine magnetotactic bacteria in a seasonally stratified coastal salt pond. Applied and Environmental Microbiology, 70(10), 6230-6239. Strickland J, P.T., (1968) A Practical Handbook of Seawater Analysis. Treusch, A.H., Leininger, S., Kletzin, A., Schuster, S.C., Klenk, H.P., Schleper, C., (2005) Novel genes for nitrite reductase and Amo-related proteins indicate a role of uncultivated mesophilic crenarchaeota in nitrogen cycling. Environmental Microbiology, 7(12), 1985-1995. Ward, B.B., O'Mullan, G.D., (2002) Worldwide distribution of Nitrosococcus oceani, a marine ammonia-oxidizing gamma-proteobacterium, detected by PCR and sequencing of 16S rRNA and amoA genes. Applied and Environmental Microbiology, 68(8), 4153-4157.

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Figure 1 Microbial nitrogen transformations above, below and across an oxic/anoxic interface in the marine environment (based in part Figure 1. on Arrigo, 2005). The marine nitrogen cycle highlighting the role of nitrification in Nitrite is highlighted in red to emphasize the central role of this metabolic intermediate/product within and between N- cycling pathways. Key functional genes discussed in the text are shown in yellow: amo, ammonia mono-oxygenase; hao, bacterial connecting the remineraliztion of particulate organic nitrogren and nitrogen loss hydroxylamine oxidoreductase (? unknown gene/enzyme in AOA); nir, nitrite reductase; and nor, nitric oxide reductase. For clarity, ¼ from the ocean. Modified from other functional genes and the process of nitrate/nitrite(Francis assimilation et al. are, 2007) not shown..

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Figure 9. Nitrogen species in the water column (left) and quantification of archaeal and bacterial amoA copy number.

!"#$%&'()*& +&(,-./)0&& 1&23&'()*&& !"#$%&$'()*+,,-".+*-/.).0."1*+'#2+(-"** 34544** 67** 8'("+'#2+(-&(*("'.#2,("&*#$%&$'(*#%-"(* 3>5?6** @** 9+"*:'+"#.;#-*<+=** !"#$%&$'()*#'("+'#2+(-&(** ?@5>A** 6?4** !"#$%&$'()*+'#2+(-"* B3536** 4** 8'("+'#2+(-&(*("'.#2,("&*#$%&$'(*#%-"(** 3?564C?A5@D** 63@**

Table 1. Top BLAST hits when from the FunGene amoA database. Legend: Sample ()*+,-,.!"#$%$/.01-$234),)5$6)--$

SH D archaeal 18 SH B archaeal 45

SH A archaeal 25 SH D archaeal 12

SH D archaeal 16

SH B archaeal 68

SH D archaeal 46

SH B archaeal 80

SH D archaeal 14 SH B archaeal 1

SH D archaeal 19

SH D archaeal 10

SH B archaeal 49

SH D archaeal 32

SH B archaeal 73

SH B archaeal 61 SH B archaeal 59 SH B archaeal 71 SH D archaeal 4

SH A archaeal 23 SH D archaeal 26 SH D archaeal 29 SH D archaeal 41

SH A archaeal 5 SH B archaeal 70 SH A archaeal 33

SH A archaeal 27 SH B archaeal 52 SH B archaeal 87 SH B archaeal 26 SH B archaeal 53 SH A archaeal 4 SH D archaeal 13 SH D archaeal 63

SH D archaeal 67 SH A archaeal 44

SH A archaeal 21 SH D archaeal 47 SH D archaeal 27

SH D archaeal 40 SH A archaeal 14

SH D archaeal 34 SH A archaeal 40 SH D archaeal 31 SH D archaeal 33 SH D archaeal 44 SH D archaeal 23 SH D archaeal 53

SH D archaeal 20 SH D archaeal 65

SH B archaeal 78 SH A archaeal 47 SH D archaeal 69 SH B archaeal 28 SH D archaeal 7 SH B archaeal 5 SH D archaeal 45

SH D archaeal 5 SH D archaeal 49

SH B archaeal 38 SH B archaeal 81

SH A archaeal 6 SH A archaeal 30

SH D archaeal 22 SH A archaeal 13 SH A archaeal 2

SH B archaeal 64

SH B archaeal 32

SH B archaeal 74

SH D archaeal 35

SH D archaeal 2

SH D archaeal 48 SH B archaeal 44 SH D archaeal 58 SH A archaeal 32 SH B archaeal 42

SH B archaeal 36 SH B archaeal 65

SH B archaeal 40 SH D archaeal 24

SH B archaeal 47 SH D archaeal 11

SH B archaeal 34 SH D archaeal 51

SH D archaeal 6 SH B archaeal 29 SH D archaeal 39 SH A archaeal 11 SH D archaeal 1 SH A archaeal 22 SH D archaeal 55 SH A archaeal 26 SH D archaeal 17 SH B archaeal 37 SH D archaeal 70 SH A archaeal 20 SH D archaeal 71

SH B archaeal 4 SH D archaeal 8

SH B archaeal 43 SH I archaeal 25

SH A archaeal 16 SH I archaeal 69

SH I archaeal 34 SH A archaeal 24 SH I archaeal 12 SH B archaeal 39 SH I archaeal 26 SH B archaeal 46 SH I archaeal 3 SH B archaeal 54 SH I archaeal 29 SH B archaeal 13 SH B archaeal 19 SH B archaeal 41 SH B archaeal 60

SH B archaeal 6 SH A archaeal 9

SH B archaeal 12 SH A archaeal 34

SH D archaeal 66 SH D archaeal 64 SH D archaeal 21 SH D archaeal 30 SH D archaeal 36 SH B archaeal 9 SH D archaeal 15 SH B archaeal 2 SH A archaeal 37 SH B archaeal 56 SH A archaeal 36 SH B archaeal 7 SH A archaeal 38 SH A archaeal 18 SH A archaeal 39 #$%$ SH B archaeal 62 SH A archaeal 48 SH D archaeal 52 SH B archaeal 27 SH D archaeal 60 SH A archaeal 12 SH D archaeal 9

SH B archaeal 66 SH D archaeal 3

SH B archaeal 11 SH A archaeal 3

SH A archaeal 8 SH A archaeal 46

SH B archaeal 17 SH D archaeal 42 #"'$%$ SH D archaeal 54

SH B archaeal 83 SH D archaeal 57

SH B archaeal 10 SH A archaeal 15 SH D archaeal 43 SH A archaeal 28 SH D archaeal 50

SH A archaeal 35 SH A archaeal 10

SH A archaeal 7 SH B archaeal 63

SH B archaeal 85 SH A archaeal 31 SH D archaeal 61 SH D archaeal 56

SH B archaeal 25 SH D archaeal 62

SH B archaeal 55 SH D archaeal 37 SH B archaeal 77 SH B archaeal 48

SH B archaeal 76

SH B archaeal 18 SH B archaeal 8 &"'$%$

SH B archaeal 69

SH B archaeal 35

SH A archaeal 41

SH B archaeal 20 SH D archaeal 25

SH B archaeal 15 SH A archaeal 1

SH B archaeal 3

SH B archaeal 79

SH A archaeal 49

SH A archaeal 17

SH B archaeal 86

SH A archaeal 42

SH A archaeal 45

SH A archaeal 43

SH A archaeal 19

SH B archaeal 57 SH B archaeal 51 !"#$%$

0.01 Legend: Sample Figure 10. Trees

SH D archaeal 18 SH B archaeal 45

SH A archaeal 25 SH D archaeal 12

SH D archaeal 16

SH B archaeal 68 SH D archaeal 46

SH B archaeal 80

SH D archaeal 14 SH B archaeal 1 SH D archaeal 19 of archaeal amoA SH D archaeal 10

SH B archaeal 49

SH D archaeal 32

SH B archaeal 73 SH B archaeal 59 SH B archaeal 61 SH B archaeal 71

SH D archaeal 4

SH A archaeal 23

SH D archaeal 26 SH D archaeal 29

SH D archaeal 41 SH B archaeal 70 SH A archaeal 5 SH A archaeal 33 SH B archaeal 52 SH A archaeal 27 functional gene SH B archaeal 87

SH B archaeal 26 SH B archaeal 53

SH A archaeal 4 SH D archaeal 13 SH D archaeal 63

SH D archaeal 67 SH A archaeal 44

SH A archaeal 21 SH D archaeal 47 SH D archaeal 27 SH D archaeal 40 SH A archaeal 14 clone libraries SH D archaeal 34 SH A archaeal 40

SH D archaeal 31 SH D archaeal 33 SH D archaeal 44 SH D archaeal 23 SH D archaeal 53 SH D archaeal 20 SH D archaeal 65 created in SH B archaeal 78 SH A archaeal 47

SH D archaeal 69 SH B archaeal 28 SH D archaeal 7 SH B archaeal 5 SH D archaeal 45 SH D archaeal 5 FastTree and SH D archaeal 49

SH B archaeal 38 SH B archaeal 81

SH A archaeal 6 SH A archaeal 30 SH D archaeal 22 SH A archaeal 13 visualized in iTOL. SH A archaeal 2

SH B archaeal 64

SH B archaeal 32 SH B archaeal 74 SH D archaeal 35

SH D archaeal 2

SH D archaeal 48 SH B archaeal 44 SH D archaeal 58 SH A archaeal 32 SH B archaeal 42

SH B archaeal 36 SH B archaeal 65

SH B archaeal 40 SH D archaeal 24

SH B archaeal 47 SH D archaeal 11

SH B archaeal 34 SH D archaeal 51 SH D archaeal 6 SH B archaeal 29 SH D archaeal 39 SH A archaeal 11 SH D archaeal 1 SH A archaeal 22 SH D archaeal 55 SH A archaeal 26 SH D archaeal 17 SH B archaeal 37 SH D archaeal 70

SH A archaeal 20 SH D archaeal 71

SH B archaeal 4 SH D archaeal 8

SH B archaeal 43 SH I archaeal 25

SH A archaeal 16 SH I archaeal 69

SH I archaeal 34 SH A archaeal 24 SH I archaeal 12 SH B archaeal 39 SH I archaeal 26 SH B archaeal 46 SH I archaeal 3 SH B archaeal 54 SH I archaeal 29 SH B archaeal 13 SH B archaeal 19

SH B archaeal 41 SH B archaeal 60

SH B archaeal 6 SH A archaeal 9

SH B archaeal 12 SH A archaeal 34

SH D archaeal 66 SH D archaeal 64 SH D archaeal 21 SH D archaeal 30 SH D archaeal 36 SH B archaeal 9 SH D archaeal 15 SH B archaeal 2 SH A archaeal 37 SH B archaeal 56 SH A archaeal 36 SH B archaeal 7 SH A archaeal 38

SH A archaeal 18 SH A archaeal 39

SH A archaeal 48 SH B archaeal 62 SH D archaeal 52 SH B archaeal 27 SH D archaeal 60 SH A archaeal 12 SH D archaeal 9

SH B archaeal 66 SH D archaeal 3

SH B archaeal 11 SH A archaeal 3

SH A archaeal 8 SH A archaeal 46

SH B archaeal 17 SH D archaeal 42 SH D archaeal 54

SH B archaeal 83 SH D archaeal 57

SH A archaeal 15 SH B archaeal 10 SH D archaeal 43 SH A archaeal 28 SH D archaeal 50

SH A archaeal 35 SH A archaeal 10 SH A archaeal 7 SH B archaeal 63 SH B archaeal 85

SH A archaeal 31 SH D archaeal 61 SH D archaeal 56 SH B archaeal 25 SH D archaeal 62 SH B archaeal 55 SH D archaeal 37 SH B archaeal 77 SH B archaeal 48

SH B archaeal 76 SH B archaeal 18 SH B archaeal 8 SH B archaeal 69

SH B archaeal 35

SH A archaeal 41

SH B archaeal 20 SH D archaeal 25

SH B archaeal 15 SH A archaeal 1

SH B archaeal 3

SH B archaeal 79

SH A archaeal 49

SH A archaeal 17

SH B archaeal 86

SH A archaeal 42

SH A archaeal 45

SH A archaeal 43

SH A archaeal 19

SH B archaeal 57

SH B archaeal 51 CARD-FISH Counts

% of DAPI Counts 0% 20% 40% 60% 80% 100%

0.5 m

% Cren 1.5m % Bacteria % Other

3.0 m

3.5 m

Figure 11. CARD-FISH counts with bacterial and crenarchaeal probes.

Figure 12. Flourescence imgage of cells displaying peanut-like morphology of marine Crenarchaeota.